I use 5 ng of vector (6200 bp) and the approprite amount of insert (40 bp) [I determined the amount with ligation calculator using 1:5 molar ratio.] Then I ligated 1,5 h in 15 ul final volume on 18 C.

I had success with annealing of other ss oligos (65-70 bp) and with ligation of these and larger fragments using these parameters and conditions. I don't know whether the problem is the wrong annealing or the wrong ligation...

Has anyone got any idea about the problem? A little help would be appreciated

If your oligos have no phosphate, then your vector must have phosophates for the ligation to succeed. How are you preparing the vector? Ideally, it would be double cut, with different RE sites at each end. When you do the ligation, you want dilute vector (5 ng is good) and equimolar amounts of your insert. This means very high dilution of your DNA, since the molecule is very very short.

My problem seems to be resolved. I used dephosphorylated vector in ligation because of the non-complete restriction efficiency to avoid self-ligated clones (There are two different RE sites in this cloning). But I didn't know that our oligos came without phosphate groups at ends. Therefore I made a new restriction digestion without dephosphorylation of the vectror. So, I got appropriate number of colonies relative to controll.